HAL Id: hal-02872797
https://hal.univ-angers.fr/hal-02872797
Submitted on 21 Dec 2020
HAL is a multi-disciplinary open access archive for the deposit and dissemination of sci- entific research documents, whether they are pub- lished or not. The documents may come from teaching and research institutions in France or abroad, or from public or private research centers.
L’archive ouverte pluridisciplinaire HAL, est destinée au dépôt et à la diffusion de documents scientifiques de niveau recherche, publiés ou non, émanant des établissements d’enseignement et de recherche français ou étrangers, des laboratoires publics ou privés.
Distributed under a Creative Commons Attribution - NoDerivatives| 4.0 International License
Charlotte Lekieffre, Thierry Jauffrais, Emmanuelle Geslin, Bruno Jesus, Joan Bernhard, et al.. Inor- ganic carbon and nitrogen assimilation in cellular compartments of a benthic kleptoplastic foraminifer.
Scientific Reports, Nature Publishing Group, 2018, 8 (1), pp.10140. �10.1038/s41598-018-28455-1�.
�hal-02872797�
www.nature.com/scientificreports
Inorganic carbon and nitrogen assimilation in cellular
compartments of a benthic kleptoplastic foraminifer
Charlotte LeKieffre
1,2, Thierry Jauffrais
2, Emmanuelle Geslin
2, Bruno Jesus
3,4, Joan M. Bernhard
5, Maria-Evangelia Giovani
1& Anders Meibom
1,6Haynesina germanica, an ubiquitous benthic foraminifer in intertidal mudflats, has the remarkable
ability to isolate, sequester, and use chloroplasts from microalgae. The photosynthetic functionality of these kleptoplasts has been demonstrated by measuring photosystem II quantum efficiency and
O2production rates, but the precise role of the kleptoplasts in foraminiferal metabolism is poorly understood. Thus, the mechanism and dynamics of C and N assimilation and translocation from the kleptoplasts to the foraminiferal host requires study. The objective of this study was to investigate, using correlated TEM and NanoSIMS imaging, the assimilation of inorganic C and N (here ammonium, NH
4+) in individuals of a kleptoplastic benthic foraminiferal species. H. germanica specimens were incubated for 20 h in artificial seawater enriched with H
13CO3− and 15NH
4+during a light/dark cycle. All specimens (n = 12) incorporated
13C into their endoplasm stored primarily in the form of lipid droplets.
A control incubation in darkness resulted in no 13
C-uptake, strongly suggesting that photosynthesis is the process dominating inorganic C assimilation. Ammonium assimilation was observed both with and without light, with diffuse
15N-enrichment throughout the cytoplasm and distinct
15N-hotspots in fibrillar vesicles, electron-opaque bodies, tubulin paracrystals, bacterial associates, and, rarely and at moderate levels, in kleptoplasts. The latter observation might indicate that the kleptoplasts are involved in N assimilation. However, the higher N assimilation observed in the foraminiferal endoplasm incubated without light suggests that another cytoplasmic pathway is dominant, at least in darkness.
This study clearly shows the advantage provided by the kleptoplasts as an additional source of carbon and provides observations of ammonium uptake by the foraminiferal cell.
Kleptoplasty is defined as the process in which a cell sequesters algal chloroplasts while discarding or digesting other algal components
1. This phenomenon is encountered in different organisms, such as sacoglossans (sea slugs)
2–4, ciliates
5, dinoflagellates
6–8, and benthic foraminifera
9,10.
Studies of benthic foraminiferal kleptoplasty have focused on shallow-water species inhabiting photic zones, especially Haynesina germanica and Elphidium spp. These studies have relied on ultrastructural observations and/or genetic analyses, which established the diatom origin of the kleptoplasts, or incubation/starvation exper- iments to define kleptoplast lifetimes and functionality once inside the foraminiferal cells
10–21. Additionally
22, showed that H. germanica and Elphidium williamsoni had a net uptake of inorganic carbon (H
14CO
3) in light, and experiments with oxygen microelectrodes demonstrated that maximal O
2production by H. germanica depended on light intensity and light history
11,23. A kleptoplastic strategy thus provides these organisms with both carbon
1Laboratory for Biological Geochemistry, School of Architecture, Civil and Environmental Engineering (ENAC), Ecole Polytechnique Fédérale de Lausanne (EPFL), 1015, Lausanne, Switzerland. 2UMR CNRS 6112 LPG-BIAF, Université d’Angers, 2 Boulevard Lavoisier, 49045, Angers, CEDEX 1, France. 3EA2160, Laboratoire Mer Molécules Santé, Université de Nantes, Nantes, France. 4BioISI – Biosystems & Integrative Sciences Institute, Campo Grande University of Lisboa, Faculty of Sciences, Lisboa, Portugal. 5Woods Hole Oceanographic Institution, Department of Geology & Geophysics, Woods Hole, MA, USA. 6Center for Advanced Surface Analysis, Institute of Earth Sciences, University of Lausanne, 1015, Lausanne, Switzerland. Correspondence and requests for materials should be addressed to C.L. (email: charlotte.lekieffre@epfl.ch)
Received: 18 April 2018 Accepted: 20 June 2018 Published: xx xx xxxx
OPEN
and a source of oxygen. Cesbron et al.
11hypothesized that kleptoplasts might constitute an additional carbon source that may provide the kleptoplastic foraminifera a substantial competitive advantage, especially during periods of impoverished nutrients. However, the extent to which kleptoplasty contributes to the carbon within the foraminiferal cell storage via photosynthetic C assimilation has not been studied yet.
Foraminiferal kleptoplasts might also be involved in uptake of inorganic N. Indeed, diatoms, from which foraminifera sequester their kleptoplasts
20,24,25, are able to assimilate ammonium through the chloroplast GS/
GOGAT (glutamate synthase and glutamine oxoglutarate aminotransferase) enzymatic pathway
26–28. Kleptoplasty is also shown in deep-sea species
9,24,25,29living in complete darkness and thus unable to perform photosynthe- sis
9,24. Among these deep-sea species, Nonionella stella maintains kleptoplasts and associated functional enzy- matic machinery, including ribulose bis-phosphate carboxylase oxygenase (RuBisCO) and phosphoenol pyruvate carboxylase (PEP carboxylase), intact for months in the dark after sampling
24. It was suggested that kleptoplasts in these deep-sea species are involved in assimilation of inorganic N
24. Therefore, a similar role in shallow klepto- plastic foraminiferal species is possible and needs to be investigated.
To date, no studies have documented the timing and distribution of assimilation and translocation of C-photosynthates or N-compounds between kleptoplasts and the foraminiferal cell. To precisely trace the C and N assimilation within the different cell compartments in the kleptoplastic foraminiferal cell, the NanoSIMS (nanos- cale secondary ion mass spectrometry, a relatively recent ultra-high resolution isotopic imaging method
30,31), was used in combination with transmission electron microscopy (TEM) and stable isotope labeling experiments
30,31. This combined approach has already been successfully applied to study assimilation, storage, and transfer of C and N in several different marine organisms
32–39, including foraminifera
40–43. Using this integrative approach, the present study had three objectives: (1) investigate the role of kleptoplasts in C-fixation, (2) investigate the transfer and distribution of photosynthetically produced organic C within the host; and (3) investigate the potential role of kleptoplasts in foraminiferal N metabolism.
Results
H. germanica specimens were incubated for 20 h in artificial seawater enriched with 2 mM NaH
13CO
3and 10 μM
15
NH
4Cl, following a light - dark cycle (Fig. 1, see details in Methods). Specimens were preserved for analysis at regular time intervals (i.e., after 4, 8, 12, and 20 hours).
TEM observations of foraminiferal cytoplasm. The cytoplasm of all specimens had well-preserved ultrastructure (Fig. 2A), as well as intact mitochondria with visible double-membranes and cristae (Fig. 2B).
Numerous small lipid droplets (diameter of ca. 500 nm), recognized by their waxy appearance, were observed in the cytoplasm (Fig. 2C), along with some larger lipid droplets ranging from 1 to 3 µm in diameter. Numerous small oval fibrillar vesicles (ca. 500 nm in length), with the fibrils arranged in parallel, and spherical to oval-shaped electron-opaque bodies (200–500 nm) were observed in the cytoplasm (Fig. 2D,E), along with occasional tubulin paracrystals identifiable due to the regular pattern of their ultrastructure revealed by high-magnification TEM imaging (Fig. 2F). In all specimens, we observed many small structures (2 to 3 µm in length) variable in shape but mainly ovoid (Fig. 2A) with the presence of numerous vacuoles within their matrix (Fig. 2G,H). Henceforth, we refer to these as “multi-vacuolar structures”.
In all observed specimens, TEM images of the endoplasm revealed well-preserved kleptoplasts with visible pyrenoids and thylakoids (Figs 2A, 3, 4). These kleptoplasts ranged in size from 2 to 10 µm in diameter. Generally, their outlines were circular to oval. They were distributed in the endoplasm with no clear pattern and often sur- rounded by an electron-lucent space between the kleptoplast membranes and the endoplasm. Some of the small lipid droplets were observed adjacent to the kleptoplast periphery. In some cases, lipid droplets were even closely associated with kleptoplast membranes (Fig. 3B).
Uptake of H
13CO
3−within foraminiferal cells. In Experiment 1, starting at t
=8 h,
13C-enrichments
were detected in all specimens. The signal was concentrated in fibrillar vesicles, electron opaque bodies, and
lipids (Figs 4, 5A,B,C). In contrast, only one specimen from the first time point (i.e. at 4 h) exhibited
13C-enriched
structures, concentrated in fibrillar vesicles and electron opaque bodies (Fig. 4). Although some lipid drop-
lets were present, they were only slightly enriched at 4 h (Figs 4 and 6). All specimens collected between 8
Figure 1. Schematic of Experiments 1 and 2, exposing H. germanica to different light conditions. Three
specimens were sampled at each indicated time point. See text for details.
www.nature.com/scientificreports/
and 20 h of incubation exhibited
13C-enrichments in the endoplasm. The
13C-enrichment (expressed in
δ13C) of electron-opaque bodies significantly increased during the light phase from ca. 40‰ at 4 h to 180‰ at 8 h (p
<0.05) and remained stable through the dark phase of the experiment, i.e. between 8 and 20 h of incuba- tion (Fig. 6B). The
13C-enrichments of the fibrillar vesicles and the lipid droplets were relatively stable during the incubation at ca. 200 to 300‰ and 60 to 150‰, respectively (Fig. 6C,D). The cytoplasm itself was slightly more enriched after 8 h of incubation than after 4 h, with averages of ca. 100‰ and 40‰, respectively (Fig. 6A).
However, the cytoplasmic enrichment did not change statistically between 8 h and 20 h (p
>0.05; Fig. 6A). No
13
C-enrichments were found in foraminifera incubated with H
13CO
3−in darkness (Experiment 2; Figs 6 and 7).
Uptake of
15NH
4+in the foraminiferal cell. All specimens of Experiment 1 exhibited detectable
15
N-enrichments. In the cytoplasm of H. germanica,
15N-enrichments significantly increased between 4 and 8 h Figure 2. TEM micrographs of the cytoplasm and organelles of Haynesina germanica. (A) Aspect of the cytoplasm in a chamber of the penultimate whorl. (B) Intact mitochondria with well-defined cristae and intact double-membranes. (C) Small lipid droplets. (D) Fibrillar vesicles. (E) Electron-opaque bodies, (F) Tubulin paracrystals; Inset: higher magnification revealing regular pattern of the paracrystal ultrastructural organization. (G,H) multi-vacuolar structures. Arrowheads: multi-vacuolar structures; c: chloroplast; eo:
electron-opaque bodies, fv: fibrillar vesicles, li: lipid droplets; m: mitochondria, tp: tubulin paracrystals, v-
vacuole. Scale bars: A: 2 µm; B, inset F: 200 nm; C–H: 500 nm.
(during the light phase), from ca. 250 to 550‰ (p < 0.05), and stabilized between 8 and 20 h (p > 0.05), i.e., during the dark phase (Fig. 6A). Similar to the observed
13C-enrichments, the
15N-signal was concentrated in electron-opaque bodies and fibrillar vesicles (Figs 4, 5A,B and 6). The
15N-enrichment of electron-opaque bod- ies was relatively stable from 4 to 12 h (between 1800–2900‰) and then decreased to less than 1000‰ at 20 h (p < 0.05 between 4 and 20 h; Fig. 6B). At the same time, the
15N-enrichment of the fibrillar vesicles increased throughout the incubation passing from ca. 425‰ at 4 h to 1320‰ after 20 h (p < 0.05). Some of these organelles were occasionally enriched in
15N but not in
13C. The tubulin paracrystals and the multi-vacuolar structures were also strongly enriched in
15N after 8 h (Fig. 5D,E). Kleptoplasts rarely exhibited
15N-enrichments, and if such enrichments were observed, they were always moderate to low (Fig. 5F).
In Experiment 2, after 8 h in darkness, the foraminifera had incorporated a much higher concentra- tion of
15NH
4+(Fig. 6) compared with Experiment 1 at any given time (p < 0.05); the cytoplasmic average
15
N-enrichment reached a value of ca. 1100‰ after 8 h of incubation in darkness. The fibrillar vesicles were also significantly more
15N-enriched during the second experiment reaching values of ca. 2400‰ (vs. maximum of ca. 1300‰ during Experiment 1). In contrast, the electron-opaque bodies exhibited
15N-enrichment compara- ble to those recorded during the first experiment, i.e. around ca. 2600‰. As in Experiment 1, the
15N isotopic signal was observed most concentrated in electron-opaque bodies, fibrillar vesicles, tubulin paracrystals, in the multi-vacuolar structures, as well as in a few kleptoplasts (Figs 6 and 7).
Discussion
All specimens exhibited mitochondria with intact cristae and double-membranes indicating that they were alive at the time of fixation
42,44. The kleptoplasts observed in our study correspond to the morphological description of
10for H. germanica collected from the Bourgneuf Bay (as in this study) and from the Wadden Sea (Mokbaai, NL). Specimens were well preserved with undamaged thylakoids and pyrenoids. The electron-lucent space that sometimes surrounded the kleptoplasts was also previously described by
10and ascribed to a possible fixation artefact. Indeed, TEM observations of the same species fixed using high pressure freezing and freeze substitution Figure 3. TEM micrographs of one chloroplast in Haynesina germanica cytoplasm. (A) Intact pyrenoid and thylakoids. (B) Higher magnification image showing two small lipid droplets in contact with the chloroplast membranes. The chloroplast membranes adjacent to the lipid vesicle are disrupted. li: lipid droplets, py:
pyrenoid, th: thylakoid. Scale bars: A: 2 µm; B: 500 nm.
www.nature.com/scientificreports/
instead of “classic” chemical fixation revealed kleptoplasts without electron-lucent space, with their membranes directly in contact with the surrounding foraminiferal cytoplasm
16,17.
The paired TEM-NanoSIMS observations allowed the visualization of inorganic C uptake (H
13CO
3−) within foraminiferal cells incubated under a light/dark cycle (Figs 4 and 6). The absence of
13C assimilation in contin- uous darkness (Experiment 2, Figs 6 and 7) and the observed production of O
2under light, as observed for H.
germanica in other studies
11,23, strongly suggests that H. germanica kleptoplasts have a functional Calvin-Benson
cycle, resulting in the production and transfer of
13C-photosynthates to the H. germanica cell. Foraminifera can
acquire C by different trophic mechanisms
45, but they are not known to actively uptake inorganic C in the absence
of either bacterial or algal symbionts or in the presence of kleptoplasts. We found no indications of the pres-
ence of prokaryotic symbiotic photosynthetic organisms and, therefore, suggest that the observed incorpora-
tion of
13C-bicarbonate is the result of photosynthesis occurring in the kleptoplasts. However, the absence of
Figure 4. Time-evolution of
13C and
15N uptake and localization within the cytoplasm of H. germanica during
Experiment 1 (light/dark incubation with H
13CO
3−and
15NH
4+). Left column: TEM micrographs. Middle and
right columns: corresponding NanoSIMS
δ13C and
δ15N images, respectively, expressed in ‰. Arrows: fibrillar
vesicles; arrowheads: electron opaque bodies; circles and li: lipid droplets, white triangles: multi-vacuolar
structures; c: chloroplast; tp: tubulin paracrystals. Scale bars: 2 µm.
13
C-enrichment inside the kleptoplasts (Figs 4 and 5) was unexpected. This absence of
13C-enrichment can be attributed to the fact that the
13C-photosynthates are quickly transported away from the kleptoplasts and thus the
13C-enrichment stay below the detection limit of the NanoSIMS. This hypothesis is supported by previous NanoSIMS studies of autotrophic
13C-exchanges in the symbiotic association between dinoflagellates and corals, where
13C-enrichments in dinoflagellate chloroplasts were systematically much lower than in other sub-cellular organelles
34. Additionally, studies on H. germanica have shown that the cellular machinery necessary for chloro- plast maintenance is unlikely to be functional
23, which could explain why, in our Experiment 1, the kleptoplasts did not accumulate
13C within their structures. In summary, our observations show that kleptoplasts in H. ger- manica are able to assimilate inorganic C and form
13C-photosynthates that are transferred to the host cell, but the kleptoplasts do not themselves become enriched in
13C (above the detection limit of the NanoSIMS).
The numerous multi-vacuolar structures observed in H. germanica (Fig. 2A,G,H) are somewhat similar to the bacteria observed in another benthic species, Globocassidulina cf. G. biora
46. The presence of numerous such Figure 5. Foraminiferal organelles enriched in
13C and/or
15N in Experiment 1 at different time points. Left column: TEM micrographs. Middle and right columns: corresponding NanoSIMS δ
13C and δ
15N images, respectively, expressed in ‰. (A) electron-opaque bodies (after 8 h of incubation), (B) fibrillar vesicles (after 12 h of incubation), (C) lipid droplets (after 8 h of incubation), (D) tubulin paracrystals (after 20 h of incubation), (E) multi-vacuolar structures (after 12 h of incubation), (F) chloroplasts (after 8 h of incubation).
Arrowheads: electron-opaque bodies; arrows: fibrillar vesicles; circles and li: lipid droplets; c: chloroplasts; white
triangles: multi-vacuolar structures; c: chloroplast; tp: tubulin paracrystals. Scale bars: 2
µm.
www.nature.com/scientificreports/
vacuoles within prokaryotic cells has been described and linked to different biological functions, such as buoy- ancy (gas vacuoles) in planktonic bacteria or nitrate vacuoles in filamentous sulfur bacteria
47,48. Thus these struc- tures could potentially be interpreted as endobionts. Is it noteworthy that they were not labeled in
13C (Fig. 5E).
Other NanoSIMS studies looking at
13C-bicarbonate assimilation in cyanobacteria, anaerobic photosynthetic bacteria or chemotrophic bacteria have shown strong bacterial
13C-enrichments
49–52. Therefore, even if the multi-vacuolar structures observed in our study were bacteria, the absence of
13C incorporation into their struc- ture suggests that they are not photosynthesizing or assimilating inorganic carbon and that thus they do not play any role in the inorganic
13C assimilation in Haynesina germanica.
Carbon was assimilated during the light phase, transferred to the foraminiferal cell, and accumulated in spe- cific organelles: electron-opaque bodies, fibrillar vesicles, and lipid droplets. The
13C-assimilation dynamics in Figure 6. Average
13C (black bars) and
15N (white bars) enrichment of (A) the cytoplasm, (B) electron-opaque bodies, (C) fibrillar vesicles and (D) lipid droplets of H. germanica (n
=3) in Experiments 1 and 2. Error bars represent one standard deviation. Red lines indicate natural variations in
13C (solid lines) and
15N (dotted lines) enrichments as measured by NanoSIMS in similar areas of unlabelled control H. germanica specimens (n
=3;
δ13
C
=0
±40‰, and
δ15N
=0
±60‰, 3σ).
electron-opaque bodies and fibrillar vesicles should be interpreted with caution, as these organelles are poorly understood
44. Indeed, we do not know their function(s) in the cell, how fast they are produced, their turn over and what triggers their production. In addition, we do not know if the
13C-enriched hotspots observed within the foraminiferal endoplasm correspond only to newly formed electron-opaque bodies and fibrillar vesicles, or if
13C-enriched material was added to pre-existing organelles; which is likely considering the high variability observed for enrichment values (Fig. 6). However, we note that none of these organelles exhibited an increase in their
13C-enrichment during the dark phase of Experiment 1, and none of the analyzed organelles showed
13
C-enrichment during Experiment 2. This strongly indicates that there are no cytoplasmic foraminiferal path- ways for inorganic carbon assimilation; i.e. carbon is assimilated only via the photosynthetic kleptoplasts.
Lipid droplets are considered to be the main C storage form in foraminifera
44. A similar accumulation process/
sequence has been observed in the symbiotic planktonic foraminifer Orbulina universa, where photosynthesis led to an assimilation of inorganic C (H
13CO
3−) stored in the form of lipid droplets
43. In kleptoplastic sea slugs (e.g., Elysia chlorotica), lipid droplets observed in the animal tissue where argued to result from the exudation of lipids from the plastids because their fatty acids had a large proportion of algal-derived eicosapentaenoic acid (20:5)
53. However, these authors could not determine whether the plastids transferred fatty acids directly via triacylglycerols (TAGs), or as free fatty acids that may be further transformed by the host into lipid droplets. The de novo produc- tion of triacylglycerol by chloroplasts in marine algae has been demonstrated
54,55. Furthermore, de novo fatty acid synthesis is known to occur in plant cell chloroplasts
56, followed by a transfer in the form of free fatty acids to the cytosol
57. Additional transfer of soluble molecules such as maltose or glucose across the chloroplast membranes through transporters also occurs in plant cells
58. The close spatial association between kleptoplast membranes and small lipid droplets observed here (Fig. 3) may indicate a potential transfer of C via exudation of small lipid droplets from kleptoplast to the H. germanica cell, although the detailed mechanisms by which the fatty acids cross kleptoplast membranes remain unknown. Unfortunately, the distribution of soluble molecules cannot be investi- gated with NanoSIMS because the sample preparation protocol causes near complete loss of such components
41.
Teugels et al.
59reported that ammonium assimilation by the kleptoplastic sacoglossan Elysia viridis was significantly higher during light exposure than in darkness. This is consistent with the glutamine oxoglutarate aminotransferase (GOGAT) enzyme pathway that requires electron donors (e.g., reduced ferredoxin) formed during photosynthetic electron transport
60. Furthermore, the glutamate synthase (GS) metabolic reaction is ATP-dependent, and gene expression of some key enzymes (GS and GOGAT) is light regulated
60. In corals, sym- biotic dinoflagellate GS/GOGAT enzymes are thought to be the main ammonium assimilation pathway
35,38,61,62. Furthermore, cnidarian cells are also known to produce cytosol glutamate dehydrogenase (GDH)
62–64, which has a dual function: 1) it converts glutamate to
α-ketoglutarate and ammonium that is subsequently assimi- lated into chloroplasts via the GOGAT pathway
59; 2) it catalyzes the opposite reaction, i.e. the amination of the
α-ketoglutarate to produce the amino acid glutamate
65.
In our study, the observation of
15N-labeled kleptoplasts in H. germanica incubated both in light and in darkness seems consistent with a GS/GOGAT kleptoplastic pathway for ammonium assimilation (Fig. 5H). However, the uptake of
15N-ammonium was higher after 8 h of incubation in total darkness (Experiment 2) than in Experiment 1 (light-dark cycle) (Fig. 6). This higher uptake in darkness compared to light is inconsistent with the light regu- lation of the GS/GOGAT enzymatic machinery
60. Ammonium incorporation might, thus, also occur by another N-assimilation pathway in foraminifera, for example, through the glutamine dehydrogenase (GDH) pathway.
Figure 7.
13C and
15N uptake and localization within the cytoplasm of H. germanica during Experiment 2
(continuous dark incubation with H
13CO
3−and
15NH
4+). Left column: TEM micrographs. Middle and right
columns: corresponding NanoSIMS
δ13C and
δ15N images, respectively, expressed in ‰. Arrows: fibrillar
vesicles; arrowheads: electron opaque dense bodies; circles: lipid droplets, white triangles: multi-vacuolar
structures; c: chloroplast, tp: tubulin paracrystals. Scale bars: 2 µm.
www.nature.com/scientificreports/
The same organelles (fibrillar vesicles, electron-opaque bodies and multi-vacuolar structures) were found to be
15N-enriched in both Experiments 1 and 2. As previously noted for
13C-assimilation, the
15N-assimilation dynamics in electron-opaque bodies and fibrillar vesicles is difficult to interpret, due to the lack of knowledge about the function of these organelles
44. However, the electron-opaque bodies and the fibrillar vesicles seem to have different patterns for
15N assimilation. While the electron-opaque bodies incorporated large amounts of
15N even after only 4 hours and then gradually lost this
15N-enrichment over time, the fibrillar vesicles assimilated
15N throughout the 20 h incubation, independent of light condition. It is noteworthy that the
15N-assimilation pattern clearly differs from the
13C-assimilation dynamics in both organelle types.
Finally, ammonium is known to be a suitable N source for many marine prokaryotes
66–68. Thus, if the multi-vacuolar structures, abundant in all H. germanica specimens (Fig. 2A,G,H) are endosymbionts (see above) they would be expected to incorporate
15NH
4+, as is indeed observed (Fig. 5G). They could, thus, constitute another putative nitrogen assimilation pathway for the benthic foraminifer H. germanica. However, we cannot conclude further in this study about the symbiotic nature of these putative prokaryotes in H. germanica, nor about their role in foraminiferal N metabolism.
A comparative study of organic C (algae) uptake through feeding between the two dominant foraminiferal species inhabiting mudflats, the akleptoplastic Ammonia sp. and kleptoplastic H. germanica, showed a higher uptake rate by the former
69. Our results highlight that H. germanica can fix inorganic carbon. Therefore, unlike Ammonia sp., H. germanica does not rely solely on heterotrophy to meet its C requirements. The mixotrophic feeding strategy of H. germanica might give a competitive advantage and allow it to become the dominant foraminifera in mudflat environments
70–72. In addition, whether H. germanica assimilates nitrogen through the kleptoplasts, potential endosymbionts, and/or by another pathway specific to foraminifera, our observations demonstrate that it is also capable of using inorganic N as a nutrient source. Further investigation is required to quantify this uptake and elucidate the role of this benthic foraminifera in the N cycle, especially since H. german- ica thrives in coastal ecosystems that are subject to increasing eutrophication
73,74.
Conclusion
Our study demonstrates inorganic C assimilation in H. germanica, most likely via the kleptoplasts. The absence of
13C assimilation in darkness combined with previous studies documenting O
2production in light strongly suggest that photosynthesis is the process dominating inorganic C-assimilation in this species. Subsequently, photosynthates are transferred to the foraminiferal cell and utilized for its metabolism. Thus, these observations clearly show the role played by the kleptoplasts in H. germanica carbon metabolism, providing the foraminiferal cell with an additional autotrophic source of C. The observation of small lipid droplets attached to the klepto- plast membranes may suggest a transfer of C from the kleptoplasts to the foraminiferal cell in the form of lipids.
However, the detailed mechanism(s) involved in this C transfer remains unknown. The kleptoplasts may also pro- vide additional N sources to foraminiferal metabolic pathways via GS-GOGAT enzymes. However, ammonium assimilation was more efficient in darkness, requiring the existence of other N-assimilation pathways.
Material and Methods
Experiment 1: light/dark cycle incubation with H
13CO
3−and
15NH
4+. Living foraminifera were collected on April 9, 2015, at low tide on the intertidal mudflat of the Bourgneuf Bay (France, 47°00
′59.4
″N 2°01
′29.8
″W). The top centimeter of the sediment was sampled, sieved over a mesh of 150
μm with in situ seawa- ter and the
>150
μm fraction was immediately transported in the dark over
∼3 hours to the laboratory in Nantes.
In the laboratory, healthy living individuals of H. germanica were selected under a binocular microscope based on their cytoplasm color (i.e. yellow-brownish material spread through all the chambers of the specimen, except the youngest chamber). The selected specimens were placed into 5 Petri dishes (5 specimens per Petri dish) filled with artificial seawater (ASW, Red Sea Salt, salinity
=35, pH
=8.0). Four of the Petri dishes contained ASW enriched with 2 mM NaH
13CO
3and 10
μM
15NH
4Cl. The fifth Petri dish contained isotopically normal artificial seawater of the same chemical composition: specimens from this dish were fixed at T0 (beginning of the experiment) and served as controls for NanoSIMS analysis (see below). All other Petri dishes were placed in an incubator (Fytoscope FS130, temperature: 18 °C, light intensity: 90
μmol m
−2s
−1). After 8 h of light exposure they were transferred to dark conditions for another 12 h. Except for the control specimens, the foraminifera remained in the spiked ASW throughout the experiment. At each time point, i.e., after 4, 8, 12, and 20 hours, one Petri dish was removed from the incubator (Fig. 1) and the 5 specimens contained in this Petri dish were immediately chemically fixed.
Experiment 2: Incubation in continuous darkness with H
13CO
3−and
15NH
4+. H. germanica speci- mens were collected on May 16, 2015, at low tide on the intertidal mudflat of the Bourgneuf Bay (France) follow- ing the same procedure as described above. Five living specimens were selected and placed in a Petri dish with artificial seawater (Red Sea Salt, salinity
=35; pH
=8.0) enriched with 2 mM NaH
13CO
3and 10
μM of
15NH
4Cl.
They were incubated in continuous darkness for 8 h (Fig. 1) and immediately chemically fixed at the end of this incubation. Control samples, which were incubated in normal seawater, where fixed at the beginning of the experiment (T0; Fig. 1).
Preparation for TEM-NanoSIMS analysis. The specimens were chemically fixed following the pro-
tocol described
44). Briefly, foraminifera were fixed immediately after removal from the incubator, with a mix
of 4% glutaraldehyde and 2% paraformaldehyde diluted in 0.1 M cacodylate buffer, 0.4 M sucrose, and 0.1 M
NaCl (pH
=7.4) at room temperature for 24 h. They were then stored at 4 °C until further processing. Further
chemical processing and transmission electron microscope (TEM) imaging of the foraminifera were performed
at the Electron Microscopy Facility of the University of Lausanne (Switzerland). After rinsing, specimens were
Foraminiferal sections were imaged with the NanoSIMS ion microprobe with a 16 keV primary ion beam of Cs
+focused to a beam spot of ca. 100–150 nm. The secondary molecular ions
12C
2−,
13C
12C
−,
12C
14N
−and
12
C
15N
−were collected simultaneously in electron multiplier detectors at a mass-resolution of ca. 10000, enough to resolve potential interferences in the mass spectrum
34,35. Isotopic image dimensions ranged from 15
×15 µm to 30
×30 µm with 256
×256 pixel resolution. For each image, 6 layers were acquired, drift corrected and super- imposed using the software L’IMAGE (developed by Dr. Larry Nittler, Carnegie Institution of Washington DC, USA). Quantified
13C/
12C and
15N/
14N ratios were obtained as follows:
C (‰) (( C
mes/ C
nat) 1) 10
13 3
δ = − ×
δ15
N (‰)
=(( N
mes/ N
nat)
−1)
×10
3where C
mesis the measured
12C
13C
−/
12C
2−ratio of the sample and C
natis the average
12C
13C
−/
12C
2−ratio measured in unlabeled samples (control). Similarly, N
mesis the measured
12C
15N
−/
12C
14N
−ratio of the sample and N
natis the average
12C
15N
−/
12C
14N
−ratio measured in unlabeled samples. The software Look@NanoSIMS
75was used to determine the isotopic enrichment of specific organelles that were identified morphologically from TEM images.
Regions of interest (ROIs) to quantify the average isotopic enrichment of the organelles were defined from the TEM images previously aligned with the NanoSIMS images (based on the
12C
14N image). For the average iso- topic enrichment of the cytoplasm, three circles of ca. 2 μm in diameter were drawn per image, avoiding highly
15
N-enriched organelles. For each specimen, between one and three NanoSIMS images were analyzed.
Statistical analysis. For each time point, three specimens were analyzed.
δ13C and
δ15N values for main orga- nelles and cytoplasm were obtained by calculating the average of ROIs within each specimen, and then calculating the average of the three specimens for each time point. The error bars provided are thus standard deviations rep- resenting inter-specimen variability. However, for the statistical analysis, a linear mixed-effects (LME) model was constructed using all the ROIs of the three specimens for each time point (taking into account pseudo-replication effects, i.e. regrouping ROIs from three different specimens into one category), followed by a Tukey multiple comparison test. The results of the Tukey multiple comparisons tests are given in the Supplementary Data S1. The statistical analyses were performed with Rstudio software using a significance level set at α
=0.05.
Data Availability. The datasets generated and/or analyzed during the current study are available in the PANGAEA Repository (https://doi.org/10.1594/PANGAEA.891407 data will be published upon acceptance of the manuscript).
References
1. Clark, K. B., Jensen, K. R. & Stirts, H. M. Survey for functional kleptoplasty among west Atlantic Ascoglossa (=Sacoglossa) (Mollusca: Opisthobranchia). The Veliger 33, 339–345 (1990).
2. Pelletreau, K. N. et al. Sea slug kleptoplasty and plastid maintenance in a metazoan. Plant Physiol. 155, 1561–1565 (2011).
3. De Vries, J., Rauch, C., Gregor, C. & Gould, S. B. A sea slug’s guide to plastid symbiosis. Acta Soc. Bot. Pol. 83, 415 (2014).
4. Serôdio, J., Cruz, S., Cartaxana, P. & Calado, R. Photophysiology of kleptoplasts: photosynthetic use of light by chloroplasts living in animal cells. Phil Trans R Soc B 369, 20130242 (2014).
5. Moeller, H. V. & Johnson, M. D. Preferential Plastid Retention by the Acquired Phototroph Mesodinium chamaeleon. J. Eukaryot.
Microbiol. 65, 148–158
6. Kim, M., Kim, S., Yih, W. & Park, M. G. The marine dinoflagellate genus Dinophysis can retain plastids of multiple algal origins at the same time. Harmful Algae 13, 105–111 (2012).
7. Nagai, S., Nitshitani, G., Tomaru, Y., Sakiyama, S. & Kamiyama, T. Predation by the toxic dinoflagellate Dinophys fortii on the ciliate Myrionecta rubra and observation of sequestration of ciliate chloroplasts. J. Phycol. 44, 909–922 (2008).
8. Nishitani, G. et al. Multiple plastids collected by the dinoflagellate Dinophysis mitra through kleptoplastidy. Appl. Environ. Microbiol.
78, 813–821 (2012).
9. Bernhard, J. M. & Bowser, S. S. Benthic foraminifera of dysoxic sediments: chloroplast sequestration and functional morphology.
Earth-Sci. Rev. 46, 149–165 (1999).
10. Jauffrais, T. et al. Ultrastructure and distribution of kleptoplasts in benthic foraminifera from shallow-water (photic) habitats. Mar.
Micropaleontol. 138, 46–62 (2018).
11. Cesbron, F. et al. Sequestered chloroplasts in the benthic foraminifer Haynesina germanica: cellular organization, oxygen fluxes and potential ecological implications. J. Foraminifer. Res. 47, 268–278 (2017).
12. Cevasco, M. E., Lechliter, S. M., Mosier, A. E. & Perez, J. Initial observations of kleptoplasty in the foraminifera of coastal South Carolina. Southeast. Nat. 14, 361–372 (2015).
13. Correia, M. J. & Lee, J. J. Fine structure of the plastids retained by the foraminifer Elphidium excavatum (Terquem). Symbiosis 32, 15–26 (2002).
www.nature.com/scientificreports/
14. Correia, M. J. & Lee, J. J. How long do the plastids retained by Elphidium excavatum (Terquem) last in their host? Symbiosis 32, 27–37 (2002).
15. Correia, M. J. & Lee, J. J. Chloroplast retention by Elphidium excavatum (Terquem). Is it a selective process? Symbiosis 29, 343–355 (2000).
16. Goldstein, S. T., Bernhard, J. M. & Richardson, E. A. Chloroplast sequestration in the foraminifer Haynesina germanica: Application of high pressure freezing and freeze substitution. Microsc. Microanal. 10, 1458–1459 (2004).
17. Goldstein, S. T. & Richardson, E. A. Fine structure of the foraminifer Haynesina germanica (Ehrenberg) and its sequestered chloroplasts. Mar. Micropaleontol. 138, 63–71 (2018).
18. Jauffrais, T., Jesus, B., Méléder, V. & Geslin, E. Functional xanthophyll cycle and pigment content of a kleptoplastic benthic foraminifer: Haynesina germanica. PLOS ONE 12, e0172678 (2017).
19. Lee, J. J. & Lanners, E. The retention of chloroplasts by the foraminifer Elphidium crispum. Symbiosis 5, 45–59 (1988).
20. Pillet, L., de Vargas, C. & Pawlowski, J. Molecular identification of sequestered diatom chloroplasts and kleptoplastidy in foraminifera. Protist 162, 394–404 (2011).
21. Pillet, L. & Pawlowski, J. Transcriptome analysis of foraminiferan Elphidium margaritaceum questions the role of gene transfer in kleptoplastidy. Mol. Biol. Evol. 30, 66–69 (2013).
22. Lopez, E. Algal chloroplasts in the protoplasm of three species of benthic foraminifera: taxonomic affinity, viability and persistence.
Mar. Biol. 53, 201–211 (1979).
23. Jauffrais, T. et al. Effect of light on photosynthetic efficiency of sequestered chloroplasts in intertidal benthic foraminifera (Haynesina germanica and Ammonia tepida). Biogeosciences 13, 2715–2726 (2016).
24. Grzymski, J., Schofield, O. M., Falkowski, P. G. & Bernhard, J. M. The function of plastids in the deep-sea benthic foraminifer.
Nonionella stella. Limnol. Oceanogr. 47, 1569–1580 (2002).
25. Tsuchiya, M. et al. Cytologic and Genetic Characteristics of Endobiotic Bacteria and Kleptoplasts of Virgulinella fragilis (Foraminifera). J. Eukaryot. Microbiol. n/a-n/a. https://doi.org/10.1111/jeu.12200 (2015).
26. Syrett, P. J. Nitrogen metabolism of microalgae. Can. Bull. Fish. Aquat. Sci. (1981).
27. Zehr, J. P., Falkowski, P. G., Fowler, J. & Capone, D. G. Coupling between ammonium uptake and incorporation in a marine diatom:
experiments with the short-lived radioisotope 13N. Limnol. Oceanogr. 33, 518–527 (1988).
28. Zehr, J. P. & Falkowski, P. G. Pathway of Ammonium Assimilation in a Marine Diatom Determined with the Radiotracer 13n1. J.
Phycol. 24, 588–591 (1988).
29. Cedhagen, T. Retention of chloroplasts and bathymetric distribution in the Sublittoral Foraminiferan Nonionellina labradorica.
Ophelia 33, 17–30 (1991).
30. Hoppe, P., Cohen, S. & Meibom, A. NanoSIMS: Technical aspects and applications in cosmochemistry and biological geochemistry.
Geostand. Geoanalytical Res. 37, 111–154 (2013).
31. Nuñez, J., Renslow, R., Cliff, J. B. & Anderton, C. R. NanoSIMS for biological applications: Current practices and analyses.
Biointerphases 13, 03B301 (2018).
32. Ceh, J. et al. Nutrient cycling in early coral life stages: Pocillopora damicornis larvae provide their algal symbiont (Symbiodinium) with nitrogen acquired from bacterial associates. Ecol. Evol. 3, 2393–2400 (2013).
33. Clode, P. L., Stern, R. A. & Marshall, A. T. Subcellular imaging of isotopically labeled carbon compounds in a biological sample by ion microprobe (NanoSIMS). Microsc. Res. Tech. 70, 220–229 (2007).
34. Kopp, C. et al. Subcellular investigation of photosynthesis-driven carbon assimilation in the symbiotic reef coral Pocillopora damicornis. mBio 6, e02299–14 (2015).
35. Kopp, C. et al. Highly dynamic cellular-level response of symbiotic coral to a sudden increase in environmental nitrogen. mBio 4, e00052–13 (2013).
36. Krueger, T. et al. Common reef-building coral in the Northern Red Sea resistant to elevated temperature and acidification. Open Sci.
4, 170038 (2017).
37. Krupke, A. et al. The effect of nutrients on carbon and nitrogen fixation by the UCYN-A–haptophyte symbiosis. ISME J. 9, 1635–1647 (2015).
38. Pernice, M. et al. A single-cell view of ammonium assimilation in coral–dinoflagellate symbiosis. ISME J. 6, 1314–1324 (2012).
39. Raina, J.-B. et al. Subcellular tracking reveals the location of dimethylsulfoniopropionate in microalgae and visualises its uptake by marine bacteria. eLife 6, e23008 (2017).
40. LeKieffre, C. et al. Surviving anoxia in marine sediments: The metabolic response of ubiquitous benthic foraminifera (Ammonia tepida). PLoS ONE 12, e0177604 (2017).
41. Nomaki, H. et al. Innovative TEM-coupled approaches to study foraminiferal cells. Mar. Micropaleontol. 138, 90–104 (2018).
42. Nomaki, H. et al. Intracellular isotope localization in Ammonia sp. (Foraminifera) of oxygen-depleted environments: Results of nitrate and sulfate labeling experiments. Front. Microbiol. 163, https://doi.org/10.3389/fmicb.2016.00163 (2016).
43. LeKieffre, C. et al. Assimilation, translocation, and utilization of carbon between photosynthetic symbiotic dinoflagellates and their planktic foraminifera host. Mar. Biol. 165 (2018).
44. LeKieffre, C. et al. An overview of cellular ultrastructure in benthic foraminifera: New observations of rotalid species in the context of existing literature. Mar. Micropaleontol. 138, 12–32 (2018).
45. Goldstein, S. T. Foraminifera: a biological overview. in Modern foraminifera (ed. Sen Gupta, B. K.) 37–55 (Springer-Verlag New- York, 1999).
46. Bernhard, J. M. Experimental and field evidence of Antarctic foraminiferal tolerance to anoxia and hydrogen sulfide. Mar.
Micropaleontol. 20, 203–213 (1993).
47. Jørgensen, B. B. & Gallardo, V. A. Thioploca spp.: filamentous sulfur bacteria with nitrate vacuoles. FEMS Microbiol. Ecol. 28, 301–313 (1999).
48. Walsby, A. E. Structure and function of gas vacuoles. Bacteriol. Rev. 36, 1–32 (1972).
49. Behrens, S. et al. Linking microbial phylogeny to metabolic activity at the single-cell level by using enhanced element labeling- catalyzed reporter deposition fluorescence in situ hybridization (EL-FISH) and NanoSIMS. Appl. Environ. Microbiol. 74, 3143–3150 (2008).
50. Finzi-Hart, J. A. et al. Fixation and fate of C and N in the cyanobacterium Trichodesmium using nanometer-scale secondary ion mass spectrometry. Proc. Natl. Acad. Sci. 106, 6345–6350 (2009).
51. Musat, N. et al. A single-cell view on the ecophysiology of anaerobic phototrophic bacteria. Proc. Natl. Acad. Sci. 105, 17861–17866 (2008).
52. Volland, J.-M. et al. NanoSIMS and tissue autoradiography reveal symbiont carbon fixation and organic carbon transfer to giant ciliate host. ISME J. 12, 714–727 (2018).
53. Pelletreau, K. N., Weber, A. P. M., Weber, K. L. & Rumpho, M. E. Lipid accumulation during the establishment of kleptoplasty in Elysia chlorotica. PLOS ONE 9, e97477 (2014).
54. Fan, J., Andre, C. & Xu, C. A chloroplast pathway for the de novo biosynthesis of triacylglycerol in Chlamydomonas reinhardtii. FEBS Lett. 585, 1985–1991 (2011).
55. Merchant, S. S., Kropat, J., Liu, B., Shaw, J. & Warakanont, J. TAG, You’re it! Chlamydomonas as a reference organism for understanding algal triacylglycerol accumulation. Curr. Opin. Biotechnol. 23, 352–363 (2012).
64. Yellowlees, D., Rees, T. A. V. & Fitt, W. K. Effect of ammonium-supplemented seawater on glutamine synthetase and glutamate dehydrogenase activities in host tissue and zooxanthellae of Pocillopora damicornis and on ammonium uptake rates of the zooxanthellae. Pac. Sci. 48, 291–295 (1994).
65. Srivastava, H. S. & Singh, R. P. Role and regulation of L-glutamate dehydrogenase activity in higher plants. Phytochemistry 26, 597–610 (1987).
66. Tupas, L. & Koike, I. Simultaneous uptake and regeneration of ammonium by mixed assemblages of heterotrophic marine bacteria.
Mar Ecol Prog Ser 70, 273–282 (1991).
67. Wheeler, P. A. & Kirchman, D. L. Utilization of inorganic and organic nitrogen by bacteria in marine systems. Limnol. Oceanogr. 31, 998–1009 (1986).
68. Zehr, J. P. & Ward, B. B. Nitrogen cycling in the ocean: New perspectives on processes and paradigms. Appl. Environ. Microbiol. 68, 1015–1024 (2002).
69. Wukovits, J., Enge, A. J., Wanek, W., Watzka, M. & Heinz, P. Effect of increased temperature on carbon and nitrogen uptake of two intertidal foraminifera Ammonia tepida and Haynesina germanica. Biogeosciences Discuss. 1–25, https://doi.org/10.5194/bg-2016- 509 (2016).
70. Cesbron, F. et al. Vertical distribution and respiration rates of benthic foraminifera: contribution to aerobic remineralization in intertidal mudflats covered by Zostera noltei meadows. Estuar. Coast. Shelf Sci. 179, 23–38 (2016).
71. Debenay, J.-P., Bicchi, E. & Goubert, E. & Armynot du Châtelet, E. Spatio-temporal distribution of benthic foraminifera in relation to estuarine dynamics (Vie estuary, Vendée, W France). Estuar. Coast. Shelf Sci. 67, 181–197 (2006).
72. Mojtahid, M. et al. Spatial distribution of living (Rose Bengal stained) benthic foraminifera in the Loire estuary (western France). J.
Sea Res. 118, 1–16 (2016).
73. Diaz, R. J. & Rosenberg, R. Spreading dead zones and consequences for marine ecosystems. Science 321, 926–929 (2008).
74. Zhang, J. et al. Natural and human-induced hypoxia and consequences for coastal areas: synthesis and future development.
Biogeosciences 7, 1443–1467 (2010).
75. Polerecky, L. et al. Look@NanoSIMS – a tool for the analysis of nanoSIMS data in environmental microbiology. Environ. Microbiol.
14, 1009–1023 (2012).